Infectivity of Hepatitis C Virus in Plasma After Drying and Storing at Room Temperature
Objective. To determine effect of environmental exposure on the survival and infectivity of hepatitis C virus (HCV).
Methods. Three aliquots of chimpanzee plasma containing HCV and proven infectious HCV inoculum were dried and stored at room temperature, 1 aliquot for 16 hours, 1 for 4 days, and 1 for 7 days. A chimpanzee (CH247) was sequentially inoculated intravenously with each of these experimental inocula, beginning with the material stored for 7 days. Each inoculation was separated by at least 18 weeks of follow‐up to monitor for infection. The concentration of HCV RNA was measured and quasi species were sequenced for each experimental inoculum and in serum samples from CH247.
Results. Evidence of HCV infection developed in CH247 only after inoculation with the material stored for 16 hours. No infection occurred after inoculation with the material stored for 7 days or 4 days. Compared with the original infectious chimpanzee plasma, the concentration of HCV RNA was 1 log lower in all 3 experimental inocula. The same predominant sequences were found in similar proportions in the original chimpanzee plasma and in the experimental inocula, as well as in serum samples from CH247.
Conclusion. HCV in plasma can survive drying and environmental exposure to room temperature for at least 16 hours, which supports the results of recent epidemiologic investigations that implicated blood‐contaminated inanimate surfaces, objects, and/or devices as reservoirs for patient‐to‐patient transmission of HCV. Healthcare professionals in all settings should review their aseptic techniques and infection control practices to ensure that they are being performed in a manner that prevents cross‐contamination from such reservoirs.
Received June 22, 2006; accepted August 21, 2006; electronically published April 16, 2007.
Hepatitis C virus (HCV) is transmitted primarily through percutaneous exposure to infectious blood, and the 2 most commonly recognized risk factors for acquiring HCV infection in the United States are receipt of blood transfusions from infectious donors and injection of illicit drugs.1,2 Since 1989, HCV testing of blood donors has virtually eliminated transfusion‐transmitted infections.3 However, infections continue to occur among injection drug users, and transmission is associated with sharing of the equipment used to administer and prepare drugs.4 HCV infections also occur among patients in healthcare settings through contamination of common injection or infusion material used by multiple patients.5 These transmission patterns are similar to those for hepatitis B virus (HBV), a virus known to be stable in the environment,6 and suggest that HCV may be transmitted directly or indirectly from contaminated surfaces, objects, or devices.
HCV RNA has been detected on environmental surfaces in settings where blood is routinely handled,7 but the ability of the virus to survive under ambient conditions has not been demonstrated. We report the results of the first study, to our knowledge, that directly determined the infectivity of HCV in plasma exposed to typical environmental conditions for up to 1 week using an experimental chimpanzee model.
Methods
Preparation of HCV Inocula
The source of the HCV used for all experimental inocula was CH910 chimpanzee plasma (second passage), genotype 1a, which contains 105 chimpanzee infectious doses (CID) per milliliter (hereafter, CH910 plasma). The plasma has been proven infectious and titrated in chimpanzees, as described elsewhere.8 Three 100‐μL aliquots of this plasma, each calculated to contain 
CID, were dried in siliconized microfuge tubes under vacuum pressure in the presence of a desiccant (Drierite; W.A. Hammond Drierite) at room temperature. Tubes were used to simulate a solid surface.6 After overnight drying (approximately 16 hours), 1 aliquot was resuspended in 300 μL of sterile water and stored at −70°C until use (hereafter, 16‐hour material). The tubes that contained the other 2 aliquots were loosely capped and transferred to a controlled environmental chamber (42% humidity, over a saturated calcium chloride solution) for 4 and 7 days, respectively, each at 25°C, then each resuspended in 300 μL of sterile water and stored at −70°C until use (hereafter, 4‐day and 7‐day material, respectively). The temperature and humidity for storage were typical of normal environmental conditions.6 Before inoculation, one‐third of each experimental aliquot was suspended in 1 mL of phosphate‐buffered saline (pH 7.6) and stored at −70°C for additional testing.
Infectivity Experiments
An HCV‐naive chimpanzee (CH247) was sequentially inoculated intravenously with each of the 3 experimental inocula, beginning with the 7‐day material. Each inoculation was separated by at least 18 weeks of follow‐up to monitor for infection. Serum samples were collected from the chimpanzee twice weekly during the follow‐up periods and tested for levels of alanine aminotransferase (ALT), HCV RNA, and antibody to HCV (anti‐HCV). Liver biopsy specimens were obtained every 1 or 2 weeks and were then subjected to histopathologic examination and tested for HCV antigen, as described elsewhere.9 The test chimpanzee was caged separately from other animals and had not been previously exposed to HCV or had any evidence of HCV infection. Housing, maintenance, and care of the chimpanzee were in compliance with or exceeded all relevant guidelines and requirements of the Animal Care and Use Committee of the Centers for Disease Control and Prevention. The clinical, virologic, and serologic profiles of 3 chimpanzees (CH1487, CH1493, and CH1555) previously infected with 3.0 × 103 CID of CH910 plasma that had not been dried or stored were included for comparison as historical controls10 (Centers for Disease Control and Prevention, unpublished data).
Detection and Quantification of HCV RNA and Testing for Anti‐HCV Antibodies
HCV RNA was quantified in all samples using a commercial polymerase chain reaction (PCR) kit (RealArt HCV LC PCR kit; Artus Biotech) in the LightCycler instrument (Roche Molecular Diagnostics), in accordance with the manufacturer’s instructions; the detection limit of the assay is 20 IU/mL. Detection of HCV RNA in serum samples from the chimpanzees was performed using a qualitative reverse‐transcriptase PCR assay (Cobas Amplicor HCV Test, version 2.0; Roche Molecular Diagnostics). Titers of HCV RNA were determined using a quantitative reverse‐transcriptase PCR kit (Cobas Amplicor HCV Monitor Test, version 2.0; Roche Molecular Diagnostics). Testing for anti‐HCV was performed by enzyme immunoassay (Ortho HCV enzyme‐linked immunosorbent assay, version 3.0; Ortho‐Clinical Diagnostics).
Cloning and Sequencing of the E1/E2 Region of the HCV Genome
To evaluate changes in the virus populations after exposure to environmental factors, we cloned and sequenced the E1/E2 region from the viral envelope of HCV in the original chimpanzee plasma samples, the dried and stored plasma samples, and the infected chimpanzee serum samples. RNA was extracted from serum or plasma by the TRIzol method (Gibco BRL), in accordance with the manufacturer's recommendations. The total RNA extracted from 100 μL of samples was reverse transcribed in a volume of 20 μL, and the resulting complementary DNA was subjected to 2 rounds of PCR. The primers used in the first round of PCR were outer sense primer HCV1278‐1306 (5′‐ATA ACG GGT CAC CGC ATG GCA TGG GAT AT‐3′) and outer antisense primer HCV1886‐1861 (5′‐CAC CAC GGG GCT GGG AGT GAA GCA AT‐3′). One tenth of the first PCR product was used in a second PCR amplification that used the following internal primers: internal sense primer HCV1284‐1312 (5′‐GGT CAC CGC ATG GCA TGG GAT ATG ATG AT‐3′) and internal antisense primer HCV1876‐1848 (5′‐CTG GGA GTG AAG CAA TAT ACC GGA CCA CA‐3′). Thermal conditions for both rounds of PCR were as follows: the first 5‐minute denaturation step was performed at 95°C, followed by 30 cycles each consisting of denaturation for 45 seconds at 95°C, annealing for 30 seconds at 55°C, and extension for 1 minute 30 seconds at 72°C, followed by a final extension for 7 minutes at 72°C. The resulting fragment of 591 base pairs comprised an E1/E2 segment that encompassed hypervariable region 1 (spanning nucleotide [nt] positions 1284–1876, as per the prototypic HCV genotype 1a sequence; GenBank accession no. M62321). The amplicons were cloned using the Topo TA Cloning Kit (Invitrogen), in accordance with the manufacturer’s recommendations. Approximately 20 clones were selected from each specimen, and the E1/E2 region was directly amplified with the same set of primers that were used in the second round of PCR. The PCR products were sequenced in both directions using BigDye Terminator 3 with an ABI 3100 genetic analyzer (Applied Biosystems). Sequences from all clones were aligned, and the number of viral variants in each sample was determined.
Results
All serum samples collected from CH247 during the 129 days after inoculation with the 7‐day material and the 136 days after inoculation with the 4‐day material were HCV RNA and anti‐HCV negative and had ALT levels in the reference range (Figure 1). After the chimpanzee was inoculated with the 16‐hour material, HCV RNA was detected in serum samples from day 7 after inoculation through day 201, reaching a peak concentration of 7.3 log IU/mL on day 120 after inoculation (Figure 1). HCV antigen–positive hepatocytes were observed from day 11 through day 201, and seroconversion to anti‐HCV positivity was observed on day 127 after inoculation. The ALT levels were elevated above the reference range from day 11 through day 201 after inoculation, with a peak level of approximately 7 times the cutoff value on day 133 after inoculation. On further follow‐up, the ALT levels declined to below the cutoff value from day 215 onward, and HCV RNA was undetectable from day 228 onward.
Figure 1. Time line showing the biochemical, serological, and virological profile of the chimpanzee (CH247) inoculated with experimental inocula dried and stored for 3 different periods (7 days, 4 days, and 16 hours). Vertical arrows, inoculation of CH247 with 3 dried and stored inocula; solid line, alanine aminotransferase (ALT) levels; dashed line, ALT cutoff value. Top: vertical bars, hepatitis C virus (HCV) RNA titers; empty squares, HCV RNA–negative results; horizontal bars, antibody to HCV (anti‐HCV) results.
The 3 historical control chimpanzees (CH1487, CH1493, and CH1555), each inoculated with CH910 plasma that had not been dried or stored, developed HCV infection with biochemical and histopathological evidence of acute hepatitis similar to that observed in CH247. HCV RNA was first detected in serum samples from 2 control animals, CH1487 and CH1493, on day 3 and in 1 control animal, CH1555, on day 7 after inoculation. The ALT levels were elevated above the cutoff from day 7 through day 79 in CH1487 and CH1493 and from day 14 through day 73 in CH1555. Seroconversion to anti‐HCV positivity occurred on day 63 after inoculation in CH1487 and CH1493 and on day 49 in CH1555. By the end of the follow‐up period, HCV RNA had become undetectable and ALT levels had declined to below the cutoff value in 2 of the 3 animals.
The concentration of HCV RNA was 7.53 log IU/mL in the original CH910 plasma and declined by 1 log to 6.22 log IU/mL in the 16‐hour material. No further decline in concentration was observed in the 4‐day material (6.26 log IU/mL) or the 7‐day material (6.23 log IU/mL) (Figure 2). Sequence analysis of multiple clones from the E1/E2 region of the viral envelope of HCV in the original CH910 plasma, in each of the 3 experimental inocula, and in serum samples from CH247 after infection with the 16‐hour material showed the presence of the same predominant sequences in similar proportions in all 5 samples (Figure 3).
Figure 2. Plot of fluorescence versus cycle numbers generated in a real‐time polymerase chain reaction assay for the original CH910 plasma and for experimental inocula dried and stored for 3 different periods (16 hours, 4 days, and 7 days). Viral RNA levels for the various HCV‐containing experimental materials were as follows: original CH910 plasma, 7.53 log IU/mL; 16‐hour material, 6.22 log IU/mL; 4‐day material, 6.26 log IU/mL; 7‐day material, 6.23 log IU/mL.
Figure 3. Alignment of the amino acid sequences of the E1/E2 viral envelope region from hepatitis C virus (HCV) in the original CH910 plasma; the experimental inocula dried and stored for 3 different periods (16 hours, 4 days, and 7 days) and the first HCV RNA–positive serum sample from the chimpanzee (CH247) after inoculation with the inoculum dried and stored for 16 hours. The numbers above each panel indicate the codon numbers. Shaded area, HCV hypervariable region 1 (HVR1).
Discussion
To understand more clearly how HCV is transmitted among people, the characteristics of the virus that affect infectivity must be determined, including its ability to remain viable in various environments for extended periods. The data presented in this study show that HCV survives drying and environmental exposure to room temperature for at least 16 hours, indicating that blood‐contaminated surfaces, objects, and devices can serve as reservoirs for HCV transmission.
The study results also show that the potential infectivity of HCV exposed to environmental factors cannot be determined by qualitative or quantitative measurements of HCV RNA. All 3 experimental inocula in our study had similar viral RNA levels, after decreasing by 1 log during the initial 16 hours of storage, but only 1 inoculum was infectious. Moreover, sequencing of multiple clones from the E1/E2 region of the viral envelope from HCV in the original CH910 plasma and the 3 experimental inocula showed that the spectrum of HCV quasi species was the same, suggesting that the loss of infectivity in the 4‐ and 7‐day material cannot be attributed to the selective transmission of viral quasi species. In addition, the chimpanzee inoculated with the 16‐hour material showed a pattern of HCV infection similar to the control animals inoculated with the CH910 plasma that had not been dried or stored. These data indicate that substantial decay of infectious virus is possible, perhaps due to loss of integrity of the viral envelope, while the concentration of viral genomic material (ie, RNA) remains stable.
The demonstration that HCV can survive after exposure to environmental conditions supports the results of epidemiologic studies that implicate cross‐contamination from inanimate surfaces, objects, or devices in HCV transmission. In the United States, most instances of HCV transmission in healthcare settings involve the contamination of multiple‐use medication vials and infusion bags through reuse of needles and syringes or the handling of blood‐contaminated items on the same surface as clean supplies.5,11‐14 The contaminated items were implicated in percutaneous transmission to patients during periods of use that ranged from 1 to 3 days.
Cross‐contamination from inanimate surfaces or objects has also been implicated in transmission of HCV among injection drug users. The sharing of drug “cookers” (ie, containers used for heating drugs into solution) and filtration cottons was significantly associated with HCV infection independent of sharing needles and syringes.4,15 Dried blood on cookers and cotton could be a source of infectious HCV and may explain why syringe and needle exchange programs have had limited impact on the prevention of HCV infection in injection drug users.
Similar results have been reported for HBV,12,16,17 and HBV has been shown to remain viable after drying and storage at room temperature for at least 7 days using methods similar to those used in this study.6 The transfer of virus by staff members to patients through use of contaminated multidose medication vials and intravenous solutions or from contaminated surfaces by their hands or gloves has been recognized for decades as a source of transmission for HBV, particularly among patients in long‐term hemodialysis centers.14 However, the risk for transmission of HBV from undiscerned exposures to contaminated items appears to be greater than that for HCV. Compared with HCV, HBV survives for a longer time in the environment and generally circulates in the blood of infected persons at concentrations that are 2‐4 log higher.18,19 These differences in virus characteristics likely account for the higher risk of HBV transmission in other settings (eg, households) where undiscerned exposures to blood‐ or serum‐derived fluids are an important source of infection.20,21
The limitations of this study include simulation of exposure of HCV to environmental factors under controlled laboratory conditions, the availability of only 1 experimental animal, and our inability to provide a more precise duration for HCV survival between 16 hours and 4 days. The laboratory conditions used to simulate exposure to environmental factors were adapted to those used for HBV. Although it would be ideal to reproduce these findings in additional animals, the well‐characterized source material, the typical course of infection that occurred in the experimental animal infected with the 16‐hour material, and the consistency of the results with epidemiologic studies contribute to the validity of the findings. In addition, a more precise delineation of the period of infectivity for HCV outside the host would not change the implications of this study’s results.
The results of recent epidemiologic studies that implicated cross‐contaminations from inanimate surfaces, objects, and devices in the transmission of HCV (and HBV) indicate that in some settings, aseptic techniques and infection control practices are not being routinely performed as recommended.5,11 The extent to which this might be occurring is unknown. Therefore, education programs for healthcare personnel, particularly those in outpatient settings, should include review of aseptic practices for injection and infusion procedures, such as always using a new needle and syringe every time fluid is withdrawn from a multidose vial and restricting medications and intravenous solutions to single patients or to centralized medication preparation rooms separate from areas where blood or other contaminated items are handled.5,11
Acknowledgments
We thank Walter W. Bond and Lynne Sehulster for their review of the protocol for drying the original material and the manuscript.
Financial support. The study was funded by internal funds from the Division of Viral Hepatitis, National Center for Infectious Diseases, Centers for Disease Control and Prevention.
Potential conflicts of interest. All authors report no conflicts of interest relevant to this article.
References
- 1. Alter MJ. Prevention of spread of hepatitis C. Hepatology 2002; 36(5 suppl 1):S93–S98.
- 2. Centers for Disease Control and Prevention. Recommendations for prevention and control of hepatitis C virus (HCV) infection and HCV‐related chronic disease. MMWR Morb Mortal Wkly Rep 1998; 47(RR‐19):1‐39.
- 3. Busch MP, Wilber JC, Johnson P, Tobler L, Evans CS. Impact of specimen handling and storage on detection of hepatitis C virus RNA. Transfusion 1992; 32:420‐425.
- 4. Hagan H, Thiede H, Weiss NS, Hopkins SG, Duchin JS, Alexander ER. Sharing of drug preparation equipment as a risk factor for hepatitis C. Am J Public Health 2001; 91:42‐46.
- 5. Williams IT, Perz JF, Bell BP. Viral hepatitis transmission in ambulatory health care settings. Clin Infect Dis 2004; 38:1592‐1598.
- 6. Bond WW, Favero MS, Petersen NJ, Gravelle CR, Ebert JW, Maynard JE. Survival of hepatitis B virus after drying and storage for one week. Lancet 1981; 1:550‐551.
- 7. Carducci A, Verani M, Casini B, et al. Detection and potential indicators of the presence of hepatitis C virus on surfaces in hospital settings. Lett Appl Microbiol 2002; 34:189‐193.
- 8. Bradley DW, Krawczynski K, Ebert JW, et al. Parenterally transmitted non‐A, non‐B hepatitis: virus‐specific antibody response patterns in hepatitis C virus‐infected chimpanzees. Gastroenterology 1990; 99:1054‐1060.
- 9. Krawczynski K, Beach MJ, Bradley DW, et al. Hepatitis C virus antigen in hepatocytes: immunomorphologic detection and identification. Gastroenterology 1992; 103:622‐629.
- 10. Krawczynski K, Alter MJ, Tankersley DL, et al. Effect of immune globulin on the prevention of experimental hepatitis C virus infection. J Infect Dis 1996; 173:822‐828.
- 11. Centers for Disease Control and Prevention. Transmission of hepatitis B and C viruses in outpatient settings—New York, Oklahoma, and Nebraska, 2000‐2002. MMWR Morb Mortal Wkly Rep 2003; 52:901‐906.
- 12. Comstock RD, Mallonee S, Fox JL, et al. A large nosocomial outbreak of hepatitis C and hepatitis B among patients receiving pain remediation treatments. Infect Control Hosp Epidemiol 2004; 25:576‐583.
- 13. Macedo de Oliveira A, White KL, Leschinsky DP, et al. An outbreak of hepatitis C virus infections among outpatients at a hematology/oncology clinic. Ann Intern Med 2005; 142:898‐902.
- 14. Centers for Disease Control and Prevention. Recommendations for preventing transmission of infections among chronic hemodialysis patients. MMWR Morb Mortal Wkly Rep 2001; 50(RR‐5):1‐43.
- 15. Hahn JA, Page‐Shafer K, Lum PJ, et al. Hepatitis C virus seroconversion among young injection drug users: relationships and risks. J Infect Dis 2002; 186:1558‐1564.
- 16. Des Jarlais DC, Diaz T, Perlis T, et al. Variability in the incidence of human immunodeficiency virus, hepatitis B virus, and hepatitis C virus infection among young injecting drug users in New York City. Am J Epidemiol 2003; 157:467‐471.
- 17. Samandari T, Malakmadze N, Balter S, et al. A large outbreak of hepatitis B virus infections associated with frequent injections at a physician’s office. Infect Control Hosp Epidemiol 2005; 26:745‐750.
- 18. Gilbert N, Corden S, Ijaz S, Grant PR, Tedder RS, Boxall EH. Comparison of commercial assays for the quantification of HBV DNA load in health care workers: calibration differences. J Virol Methods 2002; 100:37‐47.
- 19. Favero MS, Bond WW, Petersen NJ, Berquist KR, Maynard JE. Detection methods for study of the stability of hepatitis B antigen on surfaces. J Infect Dis 1974; 129:210‐212.
- 20. Nordenfelt E, Dahlquist E. HBsAg positive adopted children as a cause of intrafamilial spread of hepatitis B. Scand J Infect Dis 1978; 10:161‐163.
- 21. Petersen NJ, Barrett DH, Bond WW, et al. Hepatitis B surface antigen in saliva, impetiginous lesions, and the environment in two remote Alaskan villages. Appl Environ Microbiol 1976; 32:572‐574.
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The views expressed in this article are the authors' and do not necessarily represent those of the Centers for Disease Control and Prevention.